Mesenchymal stem cells support human vascular endothelial cells to form vascular sprouts in human platelet lysate-based matrices

During tissue regeneration, mesenchymal stem cells can support endothelial cells in the process of new vessel formation. For a functional interaction of endothelial cells with mesenchymal stem cells a vascular inductive microenvironment is required. Using a cellular model for neo-vessel formation, we could show that newly formed vascular structures emanated from the embedded aggregates, consisting of mesenchymal stem cells co-cultured with autologous human umbilical vein endothelial cells, into avascular human platelet lysate-based matrices, bridging distances up to 5 mm to join with adjacent aggregates with the same morphology forming an interconnected network. These newly formed vascular sprouts showed branch points and generated a lumen, as sign of mature vascular development. In two-dimensional culture, we detected binding of mesenchymal stem cells to laser-damaged endothelial cells under flow conditions, mimicking the dynamics in blood vessels. In conclusion, we observed that mesenchymal stem cells can support human umbilical vein endothelial cells in their vitality and functionality. In xeno-free human platelet lysate-based matrices, endothelial cells form complex vascular networks in a primarily avascular scaffold with the aid of mesenchymal stem cells, when co-cultured in three-dimensional spherical aggregates. Under dynamic conditions, representing the flow rate of venous vessel, mesenchymal stem cells preferably bind to damaged endothelial cells presumably assisting in the healing process.


Introduction
Mesenchymal stem cells (MSCs) are the stem cells of the connective tissue, showing a strong regenerative activity to counteract degenerative diseases due to their multi-lineage differentiation and self-renewal capability [1][2][3][4]. MSCs support the regeneration of the connective tissue capability of these newly formed vascular structures to generate tube-like structures and bridge distances of up to 5 mm to the neighboring spheroid with the same morphology was observed. Further, we investigated the dynamic adhesion of MSCs to damaged HUVECs in a microfluidic system applying a flow rate representing the blood flow in venous vessels. The MSCs preferably bind to the damaged HUVECs, presumably aiding in their regeneration.

Isolation, cultivation, and characterization of MSCs and HUVECs
Human placental tissues were obtained from healthy delivering women in accordance with the Austrian Hospital Act (KAG 1982) after written informed consent and the study was approved by the Ethic Commission of Lower Austria (GSl-EK-4/3122015). Amnion-derived MSCs from placental tissue were isolated and characterized with CD73-APC, CD90-FITC and CD105-PE-Cy7 (all from eBioscience, San Diego, CA) by flow cytometry (CytoFLEX XL, Beckman Coulter GmbH, Krefeld, Germany) as described previously [21] and HUVECs were isolated from the umbilical vein and characterized as published previously [24].

Generation of spherical aggregates in hanging drops
Spherical aggregates were generated by co-cultivation of 4500 MSCs and 500 HUVECs (isolated from the same donor material) on lids of petri dishes (Greiner Bio-One, Kremsmünster, Austria) using the hanging drops technology [21]. The cell number was determined using the Luna Automated Cell Counter (Logos Biosystems, South Korea). Cells were co-cultivated in a volume of 25 μI M-199 medium supplemented with 10% fetal bovine serum (Gibco, Thermo Fisher Scientific, USA), endothelial growth supplement (20 μg/ml, Becton Dickinson, USA) and heparin (10 IU/ml, Baxalta, Austria). Alternatively, MSC aggregates of 5000 MSCs were cultured in MSC-BM TM or MSC-GM TM (both from Lonza Group Ltd, Switzerland) using the same technology and monitoring of aggregate formation was performed by phase contrast microscopy (IMT2, Olympus Austria GmbH, Austria) equipped with a digital camera (DP50, Olympus). A detailed protocol is provided here dx.doi.org/10.17504/protocols.io. bp2l6138kvqe/v1.

Imaging of spherical aggregates by scanning electron microscopy
MSC aggregates were adhered on Nunc Thermanox TM coverslips (Nunc, Thermo Fisher Scientific, USA), fixed, dehydrated, mounted on conductive double side adhesive carbon tabs (Miere to Nano V.O.F., Netherlands), sputtered with gold and analyzed by a scanning electron microscope (FlexSEM, Hitachi Ltd. Corp., Japan) as previously described [21].

Vascular network formation in HPL-based matrices by confocal microscopy
Human thrombin (20 U/ml, Sigma-Aldrich, Germany) was added to 20% human platelet lysate (HPL, MacoPharma, France) in M-199 Medium supplemented with endothelial cell growth supplement (ECGS) and 500 μl of the mix was pipetted onto an ibidi μ-Dish (ibidi GmbH, Germany). After gel formation occurred, spherical aggregates consisting of MSCs and HUVECs were embedded on the gel surface at positions with 2-5 mm between the aggregates (Fig 3 scale bar). After 48 hours 500 μl of M-199 medium supplemented with 8% HPL and ECGS was added, and medium was exchanged once a week. New vessel formation was monitored twice a week using the ChemiDoc system (Bio-Rad Laboratories Inc., USA) and phase contrast microscopy (IMT-2, Olympus). After 21 days of cultivation, HPL-based gels were fixed with 4% formaldehyde overnight to guarantee gel integrity. Cells within the HPL-based gels were stained with AF 1 488 phalloidin (0.1 U/ml, Molecular Probes, USA) and HUVECs with a rabbit anti-human von Willebrand factor (vWF) mAb (1:100, 2 μg/ml, Dianova, Germany) followed by goat anti-rabbit lgG Alexa Fluor 1 594 (1:500, Jackson Laboratories, USA). Nuclei were stained with DAPI 1:1000 (Sigma-Aldrich, USA). Alternatively, before spheroid formation by hanging drops technique, MSCs were labeled with Cell Tracker TM Green CMFDA dye (1:1000 dilution) and HUVECs were labeled with Cell Tracker TM Red CMTPX dye (1:1000 dilution, both from Thermo Fisher Scientific, USA). The gels were covered with 500 μl PBS and z-stack analysis was performed using an Apochromat 10x objective and confocal microscopy (SP8, Leica, Germany).

Binding of MSCs to adherent HUVECs in flow cells of the BioFlux 1 200 device
An electro pneumatically controlled BioFlux 1 200 system (Fluxion Biosciences, lnc., USA) with microfluidic plates of 24 independent flow chambers with a dimension of 350 μm width, 1500 μm length and 70 μm height was used to investigate binding of MSCs to a HUVEC layer applying 2 dyne/cm 2 flow. Flow channels were coated with fibronectin (2.5 μg/cm 2 , Gibco, Thermo Fisher Scientific, USA) and 2 � 10 5 HUVECs in a volume of 200 μl EGM-2 medium (Lonza Group Ltd, Switzerland) were seeded and cultivated overnight under static conditions. To reach confluency of the HUVEC layer, perfusion with 2 dyne/cm 2 was applied for 24 to 48 hours. Damage to the HUVEC layer in an area of 150 μm in diameter (0.0176 mm 2 ) was introduced by a laser capture micro-dissector (LMD6, Leica Microsystems GmbH, Germany). After dead HUVECs were removed, MSCs were applied via the inlet terminal in a density of 1 � 10 5 cells/ml under 2 dyne/mm 2 flow rate. Binding of MSCs to the HUVEC layer was monitored by phase contrast microscopy and the reaction was stopped after 3 hours by fixation with 4% formaldehyde. Flow cells were washed and incubated with mouse anti-human VEcadherin (2 μg/ml clone 123413, R&D Systems, US) followed by goat anti-mouse lgG AF 1 594 (1:500, Jackson Laboratories, USA) to stain tight junctions of ECs and CD90 FITC (2.5 μg/ml, eBioscience, USA) to stain MSCs. Nuclei were stained with DAPI. Binding of MSCs to the EC layer was investigated using an LSM 700LS confocal laser scanning microscope (Carl Zeiss Microscopy GmbH, Germany) and the ZEN software program (Zeiss, Germany). A reference area of 0.1 mm 2 (elliptic area of 330 x 415.5 μm) was investigated by phase contrast microscopy to quantify MSC binding to the HUVEC layer.

Statistical analysis
The statistical analysis was performed in GraphPad Prism 7.02. Details on the sample size and performed statistical analysis are specified in the corresponding figure legends. Statistical significance is indicated in the figure � p<0.05.

MSCs generate delicate actin filaments within 3D spherical aggregates and contribute to the extracellular matrix for stability
MSCs of passage 1, expressing the mesenchymal specific markers CD73, CD90 and CD105, were used to generate spherical aggregates applying the hanging drops technology (Fig 1A and  S1 Fig). We showed that 5000 MSCs form stable 3D spherical aggregates using scanning electron microscopy and confocal microscopy ( Fig 1B and 1C). When spherical MSC aggregates were adhered to plastic surface for imaging, MSCs emanated from the aggregates and adhered as single cells (Fig 1D). Those single MSCs developed ventral stress fibers that were anchored at both sides to focal adhesions, spanning the entire cell and showed actin branching as described previously (Fig 1E) [21].
When spherical MSC aggregates were cultured in xeno-based medium, such as MSC-GM, MSCs generated collagen type I, collagen type IV, laminin, and fibronectin as major ECM components for aggregate stability (S2 Fig). The amount of individual ECM components (collagen type I, collagen type IV, laminin, and fibronectin) produced by MSCs within the 3D spherical aggregates did not change substantially during the period of 21 days of cultivation, except for Col IV (

Vascular sprout formation in HPL-based matrices induced by spherical MSC/HUVEC aggregates
To study the supportive role of MSCs in de novo vessel formation by HUVECs, we used 20% HPL-based scaffolds. Before 3D spherical aggregate formation, MSCs were labeled with

PLOS ONE
CellTracker TM Green and HUVECs with CellTracker TM Red. The aggregates were embedded in HPL-based matrices in distances of 2-5 mm. A digital documentation system was used to visualize vascular development over a period of 21 days (Fig 3). Spherical aggregates consisting only of HUVECs were instable and could not be transferred to the matrices, whereas MSCs alone formed stable spheroids (S4A Fig). For co-culture aggregates, the support of autologous MSCs was required to ensure viability of the HUVECs. HUVECs cultured by hanging drop with non-autologous MSCs showed a round morphology indicating dying cells (S4B Fig). We showed that the spatial organization of cellular outgrowths into the initially avascular translucent HPL-based matrices occurred within the first five days of cultivation (Fig 3A and  3B). These outgrowths changed their appearance to a spindle-shape cell morphology, forming cellular strands that continued to grow towards neighboring strands showing the same morphology (Fig 3C and 3D). These newly formed sprouts developed in different planes within the HPL-based matrices to form cord-like structures with a mix of labeled HUVECs and MSCs (Fig 3E and 3F). Within these newly formed sprouts, HUVECs showed nuclear elongation and occasionally formed tip cells positive for von Willebrand factor vWF (Fig 3E and 3F). Generation of vascular branch points and a lumen are important maturation steps of newly formed blood vessels during their establishment (Fig 3F). However, compared to 2D co-cultures, the HUVECs in our 3D spheroids expressed lower levels of vWF (Fig 4A and 4B, vWF p = 0.0034, n 2D = 10 n 3D = 13), indicating a decreased angiogenesis potency.

PLOS ONE
Analyzing the dynamics of the adherens junctions, we found that the cytoskeletal protein vinculin, which is associated with focal adhesion sites in the ECM, and the focal adhesion adaptor protein paxillin are higher expressed in the MSCs of spheroids compared to those in 2D cultures (Fig 4C-4F, vinculin p = 0,071 n 2D = 15 n 3D = 12, paxillin p = 0,008 n 2D = 10 n 3D = 7)

PLOS ONE
Besides, we showed that the newly formed vascular structures showed branch points and generated a lumen in our static system in the total absence of a blood circulation introducing flow (Fig 5E).
No evidence was found for a mixed endothelial/mesenchymal phenotype or cross-differentiation of MSCs to endothelial cells in our experimental setting (S5 Fig). The endothelial marker vWF could not be detected in MSCs stained with CellTracker TM Red after co-culturing with HUVECs for 21 days (S5 Fig). MSCs presumably have a scaffolding function in the 3D spheroids to ensure their stability and support the viability and proliferative function of HUVECs.
The vascular sprouts that emanated from a spherical aggregate consisting of co-cultured MSCs and HUVECs could bridge distances up to 5 mm to form an interconnected network with adjacent aggregates showing the same morphology after 10-15 days of cultivation (Fig 5).

Binding of MSCs to adherent HUVEC layer under flow
The capability of MSCs to bind HUVECs under dynamic fluid conditions mimicking the blood flow in venous vessels using a microfluidic chamber was investigated by the Bio-Flux 1 200 device that enabled multiple temperature-controlled flow assays to run in parallel. In contrast to commercial HUVECs, which are usually in passage 4-5, we used cells of passage 1. Previously, we could show that cells of higher passages showed less VE-cadherin-positive junctions and respond differently in their orientation to flow in the BioFlux 1 200 compared to HUVECs in passage 1. MSCs of passages >3 showed an altered mitochondrial morphology and the formation of stress fibers [21].
MSCs applied in the fluid phase bound HUVECs in a density of 35-40 MSCs/0.1 mm 2 when a flow rate of 2 dyne/cm 2 was applied (Fig 6A). The shear rate of 2 dyne/cm 2 represents

PLOS ONE
the physiological value in human venules with diameters ranging from 20-70 μm [25,26]. In case of cell damage induced by laser radiation to the HUVECs, MSCs bound the HUVECs with a higher frequency (Fig 6D, p = 0.0027, control n = 34, wound n = 6). Some MSCs formed long nano-tubular extrusions (Fig 6B, arrow), others directly bound to the damaged HUVECs.

Discussion
Tissue regeneration relies on neo-vessel formation, a process that requires MSCs to support ECs during initiation and development of vascular sprouts. Tissue damage with vascular injury leads to leakage of plasma components into the damaged site and the generation of an early provisional matrix consisting of fibrin-rich polymers with interspersed cross-linked fibronectin and platelets [27][28][29][30]. The clotting response and the release of platelet α-granule content into the fibrin-rich provisional matrix stop bleeding and create a basic scaffold for MSC-EC interaction [27]. Next to fibrinogen, more than 300 bioactive substances are released by activated platelets that can potentially interfere with MSC-EC communication [31]. We argued that fibrin-based matrices that incorporate the platelet secretome serve as ideal scaffolds to create new vessels close to physiological conditions [27][28][29][30]. Here, we established an HPL-based matrix to investigate new vessel formation induced by HUVECs which requires the support of MSCs. HUVECs co-cultured with autologous MSCs generated stable spherical aggregates, an

PLOS ONE
ideal tool for vascular engineering, since MSCs provide the ECM components necessary for aggregate stability, such as collagen type I, fibronectin, laminin, and collagen type IV [16,17,32]. This is of importance because the early provisional fibrin-rich matrix is replaced during the process of healing by locally produced fibronectin, laminin, and proteoglycans, generated by invading cells of the mesenchymal lineage [27,33]. We demonstrated that MSCs within spherical aggregates provide collagen type I and IV as well as fibronectin and laminin important for aggregate stability (Fig 2), while HUVECs did not add to ECM formation under these

PLOS ONE
conditions. The fibronectin assembly is driven by a complex process of cell binding, molecular extension of the protein through physical forces to expose multiple cryptic self-association sites [28,29]. In the polymerized state, fibronectin can be considered as a scaffold signaling protein favoring new vessel formation [29,30,[34][35][36][37][38]. Additionally, the MSCs within the 3D spheroids expressed higher levels of focal adhesion-associated proteins important for cell-cell contact and adhesion to the ECM (Fig 4C-4F). By inducing junctional remodeling in the 3D culture, MSCs may maintain vascular integrity during sprouting.
In this study we verified the formation of vascular sprouts originating from the inserted spherical MSC-HUVEC aggregates into the primarily avascular HPL-based matrices. When spherical MSC-HUVEC aggregates were embedded into HPL gels at distances of up to 5 mm between the aggregates, the sprouts grew towards the adjacent aggregate showing the same morphology and joined to a connected network. Interestingly, these outgrowths also formed a lumen, similar to results previously shown by Ruger et al [39].
Further, we found that MSCs bind to autologous HUVECs under dynamic conditions mimicking the blood flow in blood vessels [40]. The shear rate of 2 dyne/cm 2 represents the physiological values in human venules with diameters ranging from 20-70 μm [25]. When the perfused HUVEC layer was damaged by UV laser beam irradiation, MSCs bound preferentially to the HUVECs located at the rim of damage. To contact the damaged HUVECs, MSC generated long nano-tubular extrusions (Fig 6B and 6C).

Conclusion
Here, we provide evidence that spherical MSC/HUVEC aggregates can initiate new vessel formation and form vascular sprouts by self-induction in HPL-based matrices. These newly formed sprouts can bridge distances of up to 5 mm to join and form an interconnected network with an adjacent aggregate having the same morphology. Although we found no evidence for trans-differentiation of MSCs into ECs, characterized by vWF expression, in our 3D system [36], we could show that MSCs support ECs in the initiation process of new vessel formation in vitro. Our results give evidence for the capability of MSCs to interact actively with HUVECs in the process of vessel formation in artificial HPL-based matrices without the requirement of additional pro-vascular bioactive molecules. From the clinical view, these assays can give further information on the ability of MSCs to contribute to the regeneration of vascular defects. MSCs were stained with CellTracker TM Red and co-cultured with HUVECs in HPL-based medium for 21 days. As controls, stained MSCs and unstained HUVECs, respectively, were cultured in HPL-based medium for 21 days. After fixation, cells were stained with a rbAb specific for vWF as endothelial marker, counterstained with anti-rabbit AF488 (green), and DAPI to highlight the nuclei (blue). After 21 days of co-culture with HUVECs in HPL-based medium, no expression of vWF was detected in MSCs. (TIF)